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Method Article

An Ex Vivo Explant Model for Studying Glial Interactions in the Mouse Retina

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DOI:

10.3791/68482

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July 15th, 2025

 ,  , 
1Department of Ophthalmology, Schepens Eye Research Institute of Massachusetts Eye and Ear, Harvard Medical School

In This Article

Summary

Here, we provide a detailed methodology for isolating the retina from the mouse eye for extended ex vivo experimentation. This protocol emphasizes making this technically demanding approach accessible for researchers who would like to take advantage of the research avenues afforded by keeping retinal glia in situ in live tissue.

Abstract

The role of glia in glaucoma is an increasingly prominent research topic, but much remains unknown about how populations of these support cells - namely astrocytes and microglia - influence retinal ganglion cell survival. While in vivo and in vitro models provide a degree of insight, both approaches have significant limitations, such as the impact of peripheral immune response in the former and changes to physiological function induced by cell isolation and culture in the case of the latter. To minimize these confounding factors, we have developed an ex vivo retinal explant system in which astrocytes, microglia, and other retinal cell types can be maintained in situ for periods of at least 3 days, enabling targeted investigation at a higher throughput than is typically feasible with in vivo models. Crucially, this approach is highly amenable to methodologies that would be challenging or unfeasible in a living animal yet remains compatible with common downstream assays of intact nervous tissue. Here, we present a protocol suitable for the isolation and culturing of intact retinal explants, along with representative results of immunofluorescence microscopy documenting the changes undergone by glia and retinal ganglion cells in the ex vivo retina.

Introduction

The light sensitive retina, an extension of the central nervous system (CNS) located at the back of the eye, is essential for sight but vulnerable to both acute injury and chronic disease. As with other CNS regions, neurons in the adult retina are not replaced when lost, and the retinal ganglion cells (RGCs) that aggregate and relay visual information to the brain are particularly vulnerable to dysfunction and death in glaucoma, resulting in irreversible vision loss1,2. Glaucoma is a leading cause of blindness worldwide3, yet despite the devastating impact on affected individuals and the overall costs to society, much remains unknown about how the early stages of the disease contribute to RGC loss4. The role of glia is a major area of investigation in glaucoma pathophysiology, as these non-neuronal support cells are essential for RGC survival5 but undergo phenotypic changes in disease that may diminish beneficial behavior6 or even drive the adoption of deleterious phenotypes7. Although the term glia encompasses a range of specialized cell types throughout the CNS8, these can be broadly divided into two categories - those that share a developmental lineage with neurons and provide trophic, energetic, and structural support9, and those with a myeloid origin, which perform specialized immune surveillance and response tasks10. Representatives of both categories - astrocytes and Müller cells in the former, microglia in the latter - populate the retina5 and undergo major changes in glaucoma that raise significant questions about their role in disease progression and RGC survival11.

Efforts to answer these questions are hindered in part by intrinsic characteristics of retinal glia that present challenges to both in vivo and in vitro investigation. Unlike neurons, they are relatively silent electrically, making approaches such as ERG that enable functional assessment of RGCs and photoreceptors in vivo unsuitable for investigating changes in glial function and behavior. And while inducible12 and spontaneous13 models of glaucoma are available, much remains unknown about changes in glia at the earliest time points in the disease, which may precede detectable RGC loss, a problem compounded by the variable penetrance of disease state in many of these models. Furthermore, evidence from both clinical and experimental glaucoma suggests contributions from peripheral immune cells that can be difficult to disambiguate from those of microglia, despite indicators that they may act in different 'directions' to influence disease progression14,15.

Challenges also abound in studying retinal glia in vitro. Although isolation and culture of astrocytes16,17 and microglia18 from the brain are well established, recent work highlights extensive glial heterogeneity between CNS regions, especially in astrocytes19, and phenotypes present in one region may not be present in glia from another, such as the retina20,21. However, directly isolating retinal astrocytes and microglia for study is particularly challenging, as both cell types are relatively sparse - each making up less than 1% of the estimated 6.5 million cells in the mouse retina22,23,24. Furthermore, unlike neurons, glia are highly plastic and rapidly adapt to dramatic changes in their surroundings16,18,25; as a result, behaviors observed in these cells in vitro may represent specific adaptations to their new environment rather than phenotypic patterns that would be typically seen in health or disease. Given the limitations of both in vivo experimentation and primary culture of retinal glia, we have sought an intermediate approach - retinal explants - in which glia, RGCs, and other elements of the retina are preserved in situ in an ex vivo context. Relative to in vivo models, this approach offers an abbreviated experimental time course26, enables direct experimental manipulation of retinal glia27, and avoids the potentially confounding influence of peripheral immune cell infiltration14. Conversely, unlike primary cell culture, there is minimal disruption of the extracellular environment, and glia remain intact and morphologically recognizable, obviating the challenges associated with essentially regrowing and identifying these cells after enzymatic and mechanical disruption16,25.

Relative to previously described retinal explant methodologies, this approach emphasizes a focus on technical reliability, reproducibility, and maximizing the 'user-friendliness' of the approach to improve accessibility26,28. In personal communications with other researchers, we found that the technical challenges associated with handling the live retina present a major hurdle to many looking to utilize explants, whereas maintenance of the explanted retina in culture was relatively straightforward for groups with appropriate cell culture facilities. Therefore, this protocol includes a number of innovations intended to reduce the learning curve associated with retina isolation and allow researchers to more rapidly begin collecting experimental data. Finally, although we have placed special emphasis on the potential of this explant model for investigating the behavior of retinal glia and characterize it primarily with immunofluorescence microscopy, other retinal cell types and structures are largely conserved as well, and the explanted retinas remain amenable to a wide range of additional investigatory techniques.

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Protocol

All procedures involving animals were approved by the Institutional Animal Care and Use Committee (IACUC) at Schepens Eye Research Institute (Protocol # 2022N000030, approved 3/4/2025). Animals were handled in accordance with the Association for Research in Vision and Ophthalmology (ARVO) Statement for the Use of Animals in Ophthalmic and Vision Research.

NOTE: Figure 1 shows the key steps in the isolation of live retina from the enucleated mouse eye.

Retina extraction process diagram; step-by-step with dissection tools, shown in petri dish setup.
Figure 1: A diagrammatic representation of key steps in the isolation of live retina from the enucleated mouse eye. The corresponding step numbers of the key steps are provided in circles. Please click here to view a larger version of this figure.

1. Preparation

  1. Disinfect a 1 mL pipet and a pair of angled forceps and place them in the biosafety cabinet. These will be needed in section 3 of the protocol.
  2. Modify a transfer pipet by cutting perpendicularly across the shaft approximately 2 cm below the reservoir, where the bore is widest, to shorten the pipet and widen the opening. Ensure a clean cut on which the retina will not snag.
    NOTE: This tool can be kept for reuse if clean and rinsed with sterile water between uses.
  3. Prepare 50 ml of explant media aseptically in a biosafety cabinet by supplementing 47.5 mL of Neurobasal media with 500 µL of N-2 supplement, 1 mL of B-27 supplement, 500 µL of Glutamax, and 500 µL of penicillin-streptomycin mix (final concentration of 100 U/mL of penicillin and 100 mg/mL of streptomycin).
    NOTE: Prepare fresh explant media for each series of isolations; excess media can be kept at 4 °C for up to 1 week and used for media changes if needed. After opening, replace unused Neurobasal medium after 1 month. N-2 and B-27 should be kept frozen as single-use aliquots, avoiding freeze-thaw cycles.
  4. In a biosafety cabinet, aseptically prepare culture plate(s) by adding an insert into each well to be used, handling via the 'prongs' that emerge from the insert rim to avoid touching surfaces that will contact culture media.
    1. Add 1 mL of explant media to the insert, followed by an additional 3 mL added to the well itself. Add 2-3 mL of sterile water to any unused wells to control evaporation.
    2. Transfer plate(s) to the incubator at 37 °C with 5% CO2 to equilibrate approximately 30 min before tissue isolation.
  5. Prepare filter squares to create a substrate for tissue stabilization and transfer. Extract the filter from a bottle-top filtration kit by cutting around the periphery of the material with a razor or scalpel. Take care to avoid injury or damaging the filter through folding or tearing.
    NOTE: Removing the clear plastic reservoir using a sturdy razor to cut the more malleable opaque plastic where the threaded adapter meets the reservoir simplifies the extraction process, but additional caution should be used to prevent injury.
    1. After the filter has been detached by cutting, use forceps to extract it. Note the orientation of the filter during and after extraction; the matte side (which faces the reservoir) should be upright for mounting the retina in step 2.16 for transfer.
    2. Cut the filter into small squares approximately 7 mm × 7 mm; each retinal explant will require one square. Pre-soak the squares in sterile water in a 100 mm Petri dish for at least 20 min.
      NOTE: Filter squares can remain immersed in sterile water for up to 1 week prior to use. Due to the tendency of the filter squares to develop high levels of static charge prior to immersion, prepare only as many squares as needed. The remaining dry filter material can be kept indefinitely in a sterile 100 mm Petri dish for later use.

2. Isolation and mounting

  1. Euthanize the mouse via an approved method before confirming with a secondary approach; this study used CO2 asphyxiation followed by cervical dislocation. Enucleate both eyes with curved, blunt-tipped forceps and place them in sterile PBS.
  2. Prepare the workspace by filling a 100 mm Petri dish with sterile PBS and placing it on the stage of the binocular dissection scope.
    1. Cut a 1 cm wide strip of lab wipe and submerge it in the dish to act as a stabilizing substrate.
    2. Transfer one eye to the dissection dish, leave the other in PBS, and place on ice or in a refrigerator at 4 °C.
      NOTE: Due to the retina's sensitivity to prolonged postmortem interval, euthanizing one mouse at a time is recommended.
  3. Position and immobilize the eye. Use forceps to maneuver the eye onto the submerged lab wipe, which will aid stabilization.
    1. Identify an appropriate holding point, grasp it with angled forceps, and gently roll the eye so that it is held from the side or, ideally, from below. Ensure that the anterior-posterior axis (i.e., from the cornea to the optic nerve) is horizontal. This orientation improves visibility and minimizes pressure on the eye during dissection.
      NOTE: If possible, grasp extraorbital tissue to avoid puncturing or squeezing the eye, favoring muscle or conjunctiva over optic nerve or fat.
  4. While maintaining a hold with the angled forceps, ensure the eye is fully submerged and securely immobilized. Make an incision with the tip of the #11 scalpel parallel and approximately 0.5 mm posterior to the limbus, where the cornea transitions to the sclera.
    NOTE: This incision should be just large enough to fit the tip of one blade of the spring scissors. As the sharpness of the scalpel tip is essential to piercing the globe cleanly, change the blade after every two retinas.
    1. Insert one blade of the spring scissors inside the globe and begin cutting circumlimbally around the eye, gently repositioning the eye as needed using forceps.
      NOTE: This cut should be as straight and parallel to the limbus as possible; repositioning the eye may require putting down the scissors and using one pair of forceps with each hand. Once exposed, keep the retina submerged throughout the dissection.
  5. Once the cut has circumscribed the eye, carefully remove the anterior segment and lens, then rotate the eyecup to face upward to ease visual inspection and vitreous removal. If an extended piece of optic nerve remains attached, trim it to a shorter (1-2 mm) length to facilitate this.
  6. Inspect the retina for visible damage and the vitreal chamber for debris, such as pigmented cells from the RPE or choroid that may have entered during the removal of the anterior segment. Use angled forceps to immobilize the eyecup in this position.
    NOTE: For optimal results, debris in the vitreous chamber must be removed, and residual vitreous and adherent remnants of the ciliary body at the retinal periphery must be minimized. This can be a challenging process, and additional notes can be found in the 'Recognizing and troubleshooting mechanical injury' and 'Metabolic injury' sections of the Representative Results.
    1. Use the modified transfer pipet to flush the vitreous chamber with PBS to remove small particulate debris, avoiding air bubbles by keeping the pipet tip below the surface of the liquid and maintaining an adequate amount of PBS in the reservoir bulb.
    2. Remove larger debris with a fine watercolor brush, minimizing contact with the retinal surface. If necessary, remove more persistent debris with fine-tipped forceps; however, avoid direct contact of metal tool tips with the retina to prevent damage.
  7. After clearing visible debris, flush the vitreous chamber with PBS from the transfer pipet repeatedly (3-5 times), as described above. This will remove loosely adherent vitreous that may remain after debris removal. Use the fine brush to gently probe the chamber, particularly near the periphery if elements of the ciliary body remain. While the vitreous is optically transparent, it will act as a drag on brush fibers and can be detected in this way.
    NOTE: A thin residual layer of vitreous on the retina does not appear to negatively impact isolation or keeping the retina ex vivo, but the presence of significant amounts at the retinal periphery, often associated with residual structures from the ciliary body, can cause retinal folding and sample loss if not removed.
    1. If significant pockets of vitreous remain, use the brush to remove it by gently sweeping outward towards the periphery, ensuring that if the fibers of the brush contact the retina, they do so from a shallow angle and trail the brush, pulling rather than pushing vitreous from the chamber.
      NOTE: The removal of vitreous requires particularly careful handling to avoid retinal damage that may not be visible macroscopically; advice on recognizing and avoiding this damage is found in the 'Recognizing and troubleshooting mechanical injury' and 'Metabolic injury' sections of the Representative Results.
  8. Once the vitreous chamber has been cleared of debris and only minimal vitreous remains, gently separate the outer retina from the eye cup while continuing to stabilize the sample with angled forceps. Avoid damaging the optic nerve head and leave this structure intact to anchor the retina.
    1. Take a pair of forceps, keeping them in closed position with tips touching, and gently insert them between retina and choroid, using a pre-existing gap from prior handling if possible. Use the flat arms of the forceps, rather than the tips, to enlarge the pocket between the retina and choroid until these structures can be completely separated.
      NOTE: Emphasize lateral motions, rather than pushing the tips, to expand the gap between the retina and choroid. Gently twisting the forceps during these motions can improve separation while reducing sheer stress from friction between forceps and tissue.
  9. After separating the outer retina from the choroid, continue stabilizing the sample while using a second pair of forceps to pull the eyecup - sclera, choroid, etc. - downward. If the retina has been successfully separated, it should remain in place, tethered at the optic nerve head, while the eye cup can be held by a single pair of forceps. If the retina presses down with the eyecup, continue gently probing and separating points of connection other than the optic nerve head.
  10. With the retina anchored at the optic nerve head, continue immobilizing the tissue and use the second pair of forceps to bunch up the eye cup below the optic nerve head.
    1. Check to ensure that none of the retinal periphery is stuck in a folded-over position due to excess vitreous in the angle between the retina and any residual ciliary body. If necessary, flush the chamber with PBS using the transfer pipet to remove any additional debris, and gently use the brush to uncurl the tissue and eliminate excess vitreous, while taking care not to tug the tissue excessively.
  11. With the retina exposed from both sides, use the spring scissors to make a series of relieving cuts, spaced approximately 90° apart, extending in a straight line from the periphery towards the optic nerve head, stopping approximately 1 mm from the center. This minimizes the transection of retinal ganglion cell axons in the retina, improving the quality of cultured explants.
  12. While holding the submerged tissue in place, gently remove the lab wipe using forceps to pull it away from the tissue laterally before removing it from the dish. Avoid contact with the retina to prevent adhesion to the wipe and possible sample loss.
  13. Next, continue to hold the tissue and use the spring scissors to sever the optic nerve directly beneath the retina, then remove the remainder of the eyecup from the dish.
  14. Once the retina is isolated from the rest of the eye, fill a 35 mm Petri dish with PBS and use forceps to place a filter square at the bottom with the rougher matte side facing up. Take care to avoid creasing the filter, as it may impede the use of the filter as a substrate for moving the retina.
  15. Gently aspirate the retina with the transfer pipet and carefully transfer it to the 35 mm dish. Maneuver the retina with the brush to ensure the inner surface of the retina is upright, and the tissue rests directly above the filter square.
  16. Slowly aspirate PBS from the dish with the transfer pipet to lower the retina onto the filter. Pause, if necessary, to readjust the tissue before continuing until the retina has settled onto the filter.
    NOTE: Rapid aspiration can drive sudden lateral motion of the retina, resulting in damage or tissue loss. The entirety of the retina must rest on the square to avoid damage; however, if the retina settles in a problematic position, PBS can be added back to the dish until the retina is resuspended, and mounting can be re-attempted.
  17. Once the retina has settled onto the filter, use the brush to unroll any peripheral retina that may have folded over. Carefully balance the level of PBS so that the retina does not easily move from the filter while ensuring that enough remains so that the sample does not dry out and the brush can move smoothly without damaging the inner retina.
  18. Once again inspect the retina for debris. To avoid damage, clean the retina without direct handling using the transfer pipet to drip PBS onto the retina from roughly 1 cm above the tissue. This should flush away any remaining debris, but if too much PBS is added to the dish, the retina may float back up. If this occurs, perform aspiration again to remount the tissue.
    NOTE: 'Sham' samples can be generated by transferring retinas to a 24-well plate for fixation, blocking/permeabilization, and antibody staining as described elsewhere21. These sham controls are useful for troubleshooting mechanical injury to the retina that may occur during isolation.

3. Transfer to culture media and upkeep

  1. Close the lid of the 35 mm dish for transport to the biosafety cabinet, keeping the dish level to avoid dislodging the retina from the filter. If this occurs, remount as above.
  2. Place the closed 35 mm dish containing the retina in the biosafety cabinet, taking care not to touch any equipment or surfaces within the cabinet before disinfecting or changing gloves.
  3. Decontaminate gloves with ethanol and/or replace them, then move the 6-well plate pre-loaded with explant media from the incubator to the biosafety cabinet. Perform steps 3.4 and 3.5 within the biosafety cabinet, using the tools placed there during step 1.1.
  4. Remove the lid of the 35 mm dish containing the retina and use angled forceps to carefully lift the filter square without touching the retina directly. Remove the lid of the 6-well plate and position the filter over the center of the insert of an appropriate well, then slowly lower it into the media. The retina will detach from the filter during this step and float just below the air-liquid interface.
  5. Once the retina and filter have separated, slowly move the filter away from the retina and remove it from the media. Using the 1 mL pipet, remove 500 µL of media from the insert to ensure the retina lies flat by trapping it between the floor of the insert and the air-liquid interface.
  6. Replace the lid on the plate and return it to the incubator at 37 °C with 5% CO2, taking care to keep the retina near the center of the well. Up to 6 retinas can be kept in a single plate.
    NOTE: If keeping explants for 24 h or less, they can be left in the incubator until the end of the experiment. If kept for longer durations, 50% media changes after 1 day in vitro and on alternating days thereafter are recommended.

4. Fixation for immunostaining (optional)

  1. If performing immunostaining, refer to the following steps for fixation; while immunostaining can be performed using parameters previously detailed for flat-mounted retinas21, fixation of the explanted retina requires specific considerations.
  2. Remove cell culture inserts with explants and media from the 6-well culture plate and place them on a non-porous surface (such as the plastic lid of the culture plate) that can be disposed of properly after fixation, as the inserts are slightly porous and small amounts of fixative will pass through to this underlying surface.
  3. Gently remove culture media from inserts, then add 1 mL of 4% paraformaldehyde (PFA) to each. Let retinas fix for 15 min in a fume hood, then remove PFA and dispose of it according to institutional policy.
    CAUTION: Paraformaldehyde is hazardous and a precursor of formaldehyde, a known carcinogen. Exercise caution and perform fixation in a fume hood with suitable precautions.
  4. Wash retinas 3 times (for 5 min each) with PBS to remove PFA, then use a modified transfer pipet to transfer the retinas in PBS to individual wells in a 24-well plate for blocking, permeabilization, and immunostaining as described elsewhere21. Dispose of PFA-contaminated plastics (pipet tips, inserts, and the non-porous surface used during fixation) according to institutional policy.
    NOTE: The transfer pipet can be kept to transfer fixed and stained retinas to a slide for mounting and imaging. Although it should not have had direct contact with PFA, using different transfer pipets for handling live and fixed tissue is recommended.

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Results

At the macroscopic scale, explanted retinas remain essentially unchanged at both 1- and 3-day time points, although by the latter time they become relatively fragile, necessitating careful handling prior to fixation or other experimental endpoints. Retinas should be relatively flat, without folding (which can result in localized disruption to the diffusion of oxygen and nutrients), and should be suspended in the media between the air-liquid interface and the cell culture insert. By day 3, a faint streak may form on the b...

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Discussion

Organotypic culture32 approaches such as retinal explants harness the experimental flexibility and rapid turnaround time of cell culture while preserving much of the in situ context of in vivo studies, making them a potent avenue for investigating complex interactions between cell types. We present this protocol as an entry point for researchers who may have substantial experience with the eye and be comfortable manipulating a fixed retina but not a fresh one; therefore, we have ...

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Disclosures

The authors have nothing to disclose.

Acknowledgements

M.A.M. was supported by NIH/NEI R01EY035312, the Glaucoma Research Foundation Catalyst for a Cure Award, the Melza M. and Frank Theodore Barr Foundation, the Robert M. Sinskey, MD, Foundation, the Ruettgers Family Charitable Foundation, and the B.L. Manger Foundation. P.F.C. was supported by NIH/NEI 2T32EY007145. This work was also enabled by an NIH Core Grant for Vision Research P30 EY003176.  Figure 1 was generated with Biorender. The authors would like to thank Dr. Nasir Uddin for feedback on the manuscript and Dr. Qiurong Zhu for assistance and support.

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Materials

List of materials used in this article
NameCompanyCatalog NumberComments
#11 Scalpel bladesBard-Parker3,71,311
1 mL pipet ('Pipetman P1000')GilsonF144059M
100 mm Petri dishFalcon351029
16% ParaformaldehydeTed Pella Inc18,505Diluted to 4% for fixation
24 Well PlateFalcon353047For fixation, blocking, and antibody incubation
35 mm Petri dishFalcon351008
6-well plate and insert kitGreiner bio-one657641
Alexa Fluor 488-Donkey Anti-Rabbit IgG (H+L)Jackson ImmunoResearch Laboratories, Inc. 711-545-1521:800 dilution
Alexa Fluor 594 Donkey anti-Rat IgG (H+L) Highly Cross-AdsorbedInvitrogenA-212091:800 dilution
Alexa Fluor 594-F(ab')2 Donkey Anti-Mouse IgG (H+L) Jackson ImmunoResearch Laboratories, Inc. 715-586-1501:800 dilution
Alexa Fluor 647-Donkey Anti-Guinea Pig IgG (H+L)Jackson ImmunoResearch Laboratories, Inc. 706-605-1481:800 dilution
Alexa Fluor 647-F(ab')2 Donkey Anti-Chicken IgY (IgG) (H+L) Jackson ImmunoResearch Laboratories, Inc. 703-606-1551:800 dilution
Angled forceps ('5/45')FST / Dumont11251-35
Anti-Brn3a antibody (mouse)ChemiconMAB15851:200 dilution
Anti-CD206 antibody (rat)BioradMCA22351:200 dilution
Anti-GFAP antibody (chicken)Abcamab46741:1000 dilution
Anti-Iba1 antibody (rabbit)FUJIFILM Wako Pure Chemical Corporation019-197411:500 dilution
Anti-Tmem119 antibody (guinea pig)Synaptic Systems400 0041:500 dilution
B-27, 50xGibco (Thermofisher)17504044
Blunt curved forceps ('Extra Fine Graefe)FST11152-10
Bovine Serum AlbuminSigma-AldrichA9647Use 1% w/v for blocking and antibody incubation
Filter kit (0.2 µm aPES membrane, 150 mL Bottle Top Filter) fisherscientificFB12566508
fine brush, size 3/0princeton art & brush co.06435-1030
GlutaMaxGibco (Thermofisher)35050061
Lab wipes (Kim wipes) KIMTECH34155
N-2, 100xGibco (Thermofisher)17502048
Neurobasal-A Medium, minus phenol redGibco (Thermofisher)12349015
Normal Donkey SerumJackson ImmunoResearch Laboratories, Inc. 017-000-121Use at 10% for blocking and antibody incubation
Penicillin-StreptomycinGibco (Thermofisher)15140122
Pipet Tips, 1000 µLTipOne1126-7810
Spring Scissors ('Cohan-Vannas')FST15000-11
Sterile water ('Dnase, Rnase free')Invitrogen10977-015
Sterile-filtered PBSGibco10010-023
Straight forceps ('mini')FST / Dumont11200-14
Transfer Pipet, 5.8 mLfisherscientific13-711-9AMMD
Triton X-100Thermo Scientific ChemicalsA16046.APUse at 0.5% for blocking and antibody incubation

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Retinal Explant ModelNeuroinflammation GlaucomaRetinal Ganglion CellsAstrocyte MicrogliaImmunofluorescence MicroscopyRetinal Cell IsolationEx Vivo RetinaGFAP Expression
An Ex Vivo Explant Model for Studying Glial Interactions in the Mouse Retina
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